Dynamics of polyphosphate-accumulating bacteria in wastewater treatment plant microbial communities detected via DAPI (4',6'-diamidino-2-phenylindole) and tetracycline labeling.

Abstract

Wastewater treatment plants with enhanced biological phosphorus removal represent a state-of-the-art technology. Nevertheless, the process of phosphate removal is prone to occasional failure. One reason is the lack of knowledge about the structure and function of the bacterial communities involved. Most of the bacteria are still not cultivable, and their functions during the wastewater treatment process are therefore unknown or subject of speculation. Here, flow cytometry was used to identify bacteria capable of polyphosphate accumulation within highly diverse communities. A novel fluorescent staining technique for the quantitative detection of polyphosphate granules on the cellular level was developed. It uses the bright green fluorescence of the antibiotic tetracycline when it complexes the divalent cations acting as a countercharge in polyphosphate granules. The dynamics of cellular DNA contents and cell sizes as growth indicators were determined in parallel to detect the most active polyphosphate-accumulating individuals/subcommunities and to determine their phylogenetic affiliation upon cell sorting. Phylotypes known as polyphosphate-accumulating organisms, such as a "Candidatus Accumulibacter"-like phylotype, were found, as well as members of the genera Pseudomonas and Tetrasphaera. The new method allows fast and convenient monitoring of the growth and polyphosphate accumulation dynamics of not-yet-cultivated bacteria in wastewater bacterial communities.

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APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Apr. 2009, p. 2111–2121
0099-2240/09/$08.00?0 doi:10.1128/AEM.01540-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.
Vol. 75, No. 7
Dynamics of Polyphosphate-Accumulating Bacteria in Wastewater
Treatment Plant Microbial Communities Detected via DAPI
(4?,6?-Diamidino-2-Phenylindole) and Tetracycline Labeling?†
S. Gu ¨nther,1M. Trutnau,2S. Kleinsteuber,1G. Hause,3T. Bley,2I. Ro ¨ske,4
H. Harms,1and S. Mu ¨ller1*
Department of Environmental Microbiology, UFZ-Helmholtz Centre for Environmental Research, Permoserstrasse 15, 04318 Leipzig, Germany1;
Institute of Food Technology and Bioprocess Engineering, Dresden University of Technology, Bergstrasse 120, 01069 Dresden,
Germany2; Microscopy Unit, Biocenter of the University Halle-Wittenberg, Weinbergweg 22, 06120 Halle/Saale, Germany3; and
Institute of Microbiology, Dresden University of Technology, Zellescher Weg 20b, 01217 Dresden, Germany4
Received 8 July 2008/Accepted 15 January 2009
Wastewater treatment plants with enhanced biological phosphorus removal represent a state-of-the-art
technology. Nevertheless, the process of phosphate removal is prone to occasional failure. One reason is the
lack of knowledge about the structure and function of the bacterial communities involved. Most of the bacteria
are still not cultivable, and their functions during the wastewater treatment process are therefore unknown or
subject of speculation. Here, flow cytometry was used to identify bacteria capable of polyphosphate accumu-
lation within highly diverse communities. A novel fluorescent staining technique for the quantitative detection
of polyphosphate granules on the cellular level was developed. It uses the bright green fluorescence of the
antibiotic tetracycline when it complexes the divalent cations acting as a countercharge in polyphosphate
granules. The dynamics of cellular DNA contents and cell sizes as growth indicators were determined in
parallel to detect the most active polyphosphate-accumulating individuals/subcommunities and to determine
their phylogenetic affiliation upon cell sorting. Phylotypes known as polyphosphate-accumulating organisms,
such as a “Candidatus Accumulibacter”-like phylotype, were found, as well as members of the genera Pseudo-
monas and Tetrasphaera. The new method allows fast and convenient monitoring of the growth and polyphos-
phate accumulation dynamics of not-yet-cultivated bacteria in wastewater bacterial communities.
Enhanced biological phosphorus removal (EBPR) is a
widely implemented technique for phosphate removal in
wastewater treatment processes, since it is economical while
being more environmentally friendly than the traditional
chemical treatment (10). Unfortunately, despite several years
of intensive research into phosphorus-accumulating organisms
(PAOs), the EBPR process is unstable. Moreover, many pure
cultures, e.g., members of the genera Acinetobacter, Microlu-
natus, Tetrasphaera, and Lampropedia, have been isolated,
which showed traits expected from PAOs, but none of them
were found to have great significance in wastewater treatment
plants (WWTPs) (8, 33, 35, 44). Our knowledge of PAOs that
are active in WWTPs is therefore cursory at best, and it has
been suggested that bacterial species which are not yet culti-
vated may be responsible for the polyphosphate accumulation
(16). Monitoring of these organisms thus requires cultivation-
independent techniques directed at specific characteristics of
PAOs, such as the accumulated polyphosphate. For the selec-
tive analysis of PAOs within complex communities, single-cell-
based methods such as fluorescence microscopy and flow cy-
tometry appear to be most appropriate.
Polyphosphate granules were first identified in Saccharomy-
ces cerevisiae in 1888 (24). Since then, the ability to store
polyphosphates as granules within the cell was found to be
widespread among microorganisms. The inert polymers fulfill
various functions (for a review, see reference 21). They consist
of linear orthophosphate chains ranging from few to several
hundred residues which are linked by energy-rich phosphoan-
hydride bonds. As polyanions, the polyphosphate chains need
counterions to neutralize the negative charge. The most im-
portant counterions are Mg2?, Ca2?, and K?. Several other
cations, such as Mn2?, Al3?, and Fe3?, were found to be
incorporated to a lesser extent, depending on the growth con-
ditions and microorganisms under investigation (42). Cur-
rently, polyphosphate accumulation is studied by bulk mea-
surements such as
(31) or even more frequently by quantifying the activity of the
polyphosphate kinase (Ppk) (4). Single-cell approaches also
have been applied for a long time, with examples being phase-
contrast or bright-field microscopy. Neisser’s staining and
Loeffler’s methylene blue are the traditional nonfluorescent
stains for microscopic analysis of PAOs (34). Electron micro-
scopic analysis made use of labeled antibodies to facilitate the
detection of phosphate granules (32). Another compound that
is widely used to visualize polyphosphate granules is the fluo-
rescent dye 4?,6?-diamidino-2-phenylindole (DAPI) (41),
which stains cells with polyphosphate contents higher than 400
?mol g?1(dry weight) when applied at a concentration of at
least 18 ?M (5 to 50 ?g ml?1) *18, 40). DAPI staining depends
on a polyphosphate-mediated metachromatic reaction which
31P nuclear magnetic resonance analysis
* Corresponding author. Mailing address: UFZ-Helmholtz Centre
for Environmental Research Leipzig-Halle, Department of Environ-
mental Microbiology, Permoserstrasse 15, 04318 Leipzig, Germany.
Phone: 49 341 235 1318. Fax: 49 341 235 1351. E-mail: susann.mueller
@ufz.de.
† Supplemental material for this article may be found at http://aem
.asm.org/.
?Published ahead of print on 30 January 2009.
2111

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causes a shift in the emitted fluorescence from blue to a bright
yellow-green. When DAPI is applied at lower concentrations
(0.24 to 5 ?M), the resulting blue fluorescence is related to
bacterial DNA. Unfortunately, unspecific fluorescence of other
cellular constituents such as lipids also was reported when
DAPI was applied at high concentrations (180 ?M) (40). An-
other, but not commonly used, fluorescent dye is 9-aminoacri-
dine, which has properties similar to those of DAPI as it emits
blue fluorescence when binding to DNA and green fluores-
cence when binding to polyphosphate granules (21).
Individual-based fluorescent techniques can be used to de-
tect bacteria in natural communities which are able to synthesize
polyphosphates but cannot be grown in pure culture. Particu-
larly promising is the recognition of PAOs by flow cytometry in
combination with cell sorting for further phylogenetic identi-
fication, an approach to be presented in this report. A few
applications of flow cytometry to determine polyphosphate
granules have been reported. They involved DAPI alone (46)
or in combination with fluorescence in situ hybridization
(FISH) (16). However, the aforementioned unspecific DAPI
staining may spoil the quantitative and possibly even the qual-
itative determination of PAOs in natural communities.
The fluorescent antibiotic tetracycline (TC) and its deriva-
tives are frequently used in medicine as a label for calcium
deposition in bone or teeth and as a marker for membrane-
associated divalent cations (12). The binding mode is still un-
clear, although several mechanisms have been discussed (for a
review, see reference 27). When TC is bound to diamagnetic
divalent cations such as calcium and magnesium, its fluores-
cence intensity is enhanced, with excitation and emission max-
ima at 390 nm and 515 nm, respectively (at pH 7.5) (23). Using
this effect, we developed a new fluorescence technique that
makes use of the interaction of TC hydrochloride with coun-
tercations of polyphosphate granules and can be used for the
quantitative determination of polyphosphate in individual bac-
terial cells.
Here we present the development of a dual polyphosphate/
DNA fluorescent staining approach and its application to fol-
low PAO dynamics within natural communities. Representa-
tive PAOs were isolated by cell sorting and identified by
sequencing and terminal restriction fragment length polymor-
phism (T-RFLP) profiling of 16S rRNA genes. This mining of
not-yet-cultivable PAOs from wastewater samples greatly in-
creases our knowledge of genuine polyphosphate accumulators
in an EBPR process and will open options for knowledge-
based optimization of WWTPs.
MATERIALS AND METHODS
Bacterial strains and culture conditions. The bacterial strains used in this
study are shown in Table 1. Cells were cultivated in 500-ml shake flasks with
tightly closed screw caps at 150 rpm and 30°C in 100 ml of the synthetic waste-
water described by Hrenovic et al. (14). They were subjected to alternating
aerobic and anaerobic conditions (changing every 12 h). For every new anaerobic
process interval, the medium was replenished with 0.26 mM propionate, 0.1 g
liter?1peptone, and 54 ?M CaCl2as well as 83 ?M MgSO4and 1.6 mM
KH2PO4. To establish aerobic and anaerobic conditions, the culture was flushed
with sterile air and with sterile nitrogen of highest purity, respectively, for at least
60 min.
Escherichia coli and the non-polyphosphate-producing Micropruina glycogenica
(37) were grown aerobically in 300-ml shake flasks in 50 ml peptone medium
containing (per liter) 5 g peptone from meat (pancreatic), 3 g NaCl, 2 g K2HPO4,
10 g meat extract, 10 g yeast extract, and 5 g glucose (for E. coli) and in DSM 776
for M. glycogenica. Methylobacterium rhodesianum was grown aerobically in
100-ml shake flasks in 50 ml of the standard medium described by Ackermann et
al. (1). This strain was reported to produce poly-?-hydroxybutyrate (PHB) to up
to 95% of its dry weight. Methanol (120 mM) was added as a carbon and energy
source. The three control strains were cultivated at 30°C, 150 rpm, and pH 7.0.
Cultures were checked regularly for purity by streaking aliquots on agar plates of
the respective medium (given above) as well as microscopic observation.
Reactor operation. Activated sludge was harvested from one aeration tank of
the Elsterwerda WWTP (Brandenburg, Germany) and cultivated in a 7-liter
laboratory bioreactor with a working volume of 5 liters (New Brunswick BioFlow
III; New Brunswick Scientific) (see Fig. S1 in the supplemental material) at 300
rpm (except for the settling period) and 20°C. The pH value of 7 ? 0.05 was
automatically regulated by titration with either 0.6 M HCl or 0.5 M NaOH. The
reactor was operated as sequenced-batch reactor (SBR) with a cycle time of 6 h
four times a day, consisting of an aerobic period (2 h, aerated) followed by a
settling and refilling period for restoring anaerobic conditions (2 h, manual feed)
and an anaerobic period (2 h). The initial mixed-liquor suspended solids (MLSS)
were set to 2.5 g liter?1by appropriate dilution with cultivation medium (OECD
synthetic wastewater) containing (per liter) 0.16 g peptone from meat (pancre-
atic), 0.11 g meat extract, 0.03 g urea, 0.028 g KH2PO4, 0.007 g NaCl, 0.004 g
CaCl2? 2H2O, and 0.002 g MgSO4? 7H2O. Trace elements (2 ml liter?1) were
added as a solution containing (per liter) 0.07 g ZnCl2, 0.1 g MnCl2? 4H2O,
0.2 g CoCl2? 6H2O, 0.1 g NiCl2? 6H2O, 0.02 g CuCl2? 2H2O, 0.05 g
NaMoO4? 2H2O, 0.026 g Na2SeO3? 5H2O, and 1 ml 25% HCl. For overnight
SBR operation, settling and refilling were replaced by automated feeding of
concentrated medium (43 ml/6 h) to result in a final concentration equal to that
from the manual feed. The concentrations of phosphate and other components
of the medium were changed during the cultivation as indicated according to the
demands of the experiments. The oxygen was recorded by an oxygen sensor
(Applicon, The Netherlands). The pH was analyzed with a sensor from Mettler-
Toledo (Switzerland). Inorganic phosphate and nitrate were measured by ion
chromatography (IC) (Dionex Corporation). IC analysis was performed on a DX
320 IC with an EG40 eluent generator and an IonPac NG1 guard column (4 by
35 mm) connected to an IonPac AS15 (4 mm) separation column. The separation
was obtained at a flow rate of 0.4 ml min?1with a gradient program (KOH and
water) rising from 5 mM to 30 mM in the first 7 min and to 45 mM in the next
10 min. The concentration of KOH then was kept constant for 5 min and finally
decreased to 5 mM within 2 min. MLSS values (g total suspended solids liter?1),
mixed-liquor volatile suspended solids (MLVSS) values (g volatile suspended
TABLE 1. Bacterial species used in this study
OrganismSource Rationale for usage
Microlunatus phosphovorus NM-1
(DSM 10555)
Pseudomonas sp.
German Collection of Microorganisms and Cell
Cultures
Wastewater isolate (M. Eschenhagen, Dresden
University of Technology, Germany)
Wastewater isolate (M. Eschenhagen, Dresden
University of Technology, Germany)
UFZ strain collection
UFZ strain collection
German Collection of Microorganisms and Cell
Cultures
Model organism for polyphosphate
production
Model organism for polyphosphate
production
Model organism for polyphosphate
production
Negative control (DNA binding of TC)
Negative control (PHB binding of TC)
Negative control (glycogen binding of TC)
Paracoccus sp.
Escherichia coli K-12
Methylobacterium rhodesianum MB126
Micropruina glycogenica Lg2T
(DSM 15918)
2112GU ¨NTHER ET AL.APPL. ENVIRON. MICROBIOL.

Page 3

solids liter?1), chemical oxygen demand (COD) values (mg liter?1) (7), and ash
contents (g liter?1) were determined using standard methods (9a).
Cell preparation. Cells of Microlunatus phosphovorus, a Pseudomonas sp., and
a Paracoccus sp. were harvested according to the demands of the experiments. E.
coli, M. glycogenica, and M. rhodesianum were harvested at the stationary phase
of growth. The samples were washed twice with phosphate-buffered saline (PBS)
(0.4 M Na2HPO4/NaH2PO4, 150 mM NaCl, pH 7.2) to remove any disturbing
substances (cell debris or organic material) by centrifugation at 3,200 ? g for 5
min (10 min for activated sludge samples) and conserved in fixation buffer (pH
7.0) containing 5 mM BaCl2(BaCl2? 2H2O; Laborchemie Apolda, Germany), 5
mM NiCl2(NiCl2? 6H2O; Merck, Germany), and 10% sodium azide (Merck,
Germany) dissolved in PBS (1 ml fixation buffer for approximately 3 ? 108cells
ml?1) as described by Gu ¨nther et al. (11) for a maximum of 9 days. For further
information on cell preparation, see reference 11.
Staining procedures and flow cytometry. (i) DNA. DNA staining was done as
described previously (11), using a variation of a standard procedure (25). A
DAPI solution of 0.24 ?M was used for DNA staining of activated sludge
samples, and a 1 ?M solution was used for M. phosphovorus.
(ii) Polyphosphates. For cellular polyphosphate staining, two different dyes
were applied. The first one was DAPI, which is known to stain polyphosphates at
high concentrations. Two milliliters of adjusted cell suspension was centrifuged
and treated with 1 ml solution A for 20 min. The cells were then washed and
resuspended carefully in 2 ml of a 28 ?M DAPI stock solution.
The second dye was the antibiotic TC hydrochloride (Fluka, Switzerland). It
was applied as a 2-mg ml?1stock solution in double-distilled water to give a final
concentration of 0.225 mM in the cell suspension. Stock solutions were prepared
freshly before each experiment to prevent low data quality due to TC aging. The
dramatic decline in fluorescence intensity of the TC stock solution after storage
for only 3 days in a refrigerator is presented in the inset of Fig. S2 in the
supplemental material. The stained samples were stored for 60 min at 20°C in the
dark before flow cytometric measurement. For the experiments with activated
sludge, unstained cells served as controls. The number of cells displaying strong
green autofluorescence was subtracted from the fluorescence counts obtained
from TC-stained samples to obtain polyphosphate-related fluorescence informa-
tion (see also the description of the gating strategy below).
(iii) DNA and polyphosphates. For dual staining of the DNA and polyphos-
phate granules by DAPI and TC, the cells were first subjected to solution A for
20 min, centrifuged, washed, and treated with 0.225 mM TC for 10 min before 2
ml solution B (0.24 ?M and 1 ?M for activated sludge and M. phosphovorus,
respectively) was added. Stained samples were incubated for another 60 min at
20°C in the dark before flow cytometric measurement.
(iv) PHB. To visualize cellular PHB contents, Nile red (Sigma) was applied as
described previously (1).
(v) Scatter behavior. Forward scatter (FSC) is related to cell size, and side
scatter (SSC) is related to cell granularity. Data were obtained by examining the
light-scattering behavior of individual cells, mediated by the 488-nm line of the
argon ion laser. Usually, shifts in the FSC/SSC mean values were recorded in
parallel with the fluorescence measurements.
(vi) Flow cytometry. Analyses were carried out using a MoFlo cell sorter
(DakoCytomation) equipped with two water-cooled argon-ion lasers (Innova
90C and Innova 70C from Coherent). Excitation of 400 mW at 488 nm was used
to analyze the Nile red fluorescence, FSC, and SSC at the first observation point.
SSC was used as a trigger signal to discriminate bacterial cells from electronic
noise. DAPI and TC were excited by 100 mW of multiline ultraviolet (333 to 365
nm) at the second observation point. The orthogonal signal was first reflected by
a beam splitter and then recorded after reflection by a 555-nm long-pass dichroic
mirror and passage by a 505-nm short-pass dichroic mirror and a 488/10 band-
pass (BP) filter. Blue (DAPI) fluorescence was passed through a 450/65 BP filter,
green (TC and DAPI) fluorescence through a 520/15 BP filter, and red (Nile red)
fluorescence through a 620/45 BP filter. Photomultiplier tubes were obtained
from Hamamatsu Photonics (models R 928 and R 3896; Hamamatsu, Japan).
Amplification was carried out at logarithmic scales for all measurements. Tuning
of the device was done as described by Kleinsteuber et al. (19).
Sorting of TC-stained, polyphosphate-containing bacterial cells was done using
the most accurate sort mode (single and one-drop mode; highest purity, 99%) at
a rate not higher than 1,500 cells per second. Cell sorting was performed using
the four-way sort option at high speed (12 ms?1). The cells were sorted into
nucleic acid-free glass flasks. Cells were separated from the mixed culture using
TC-polyphosphate fluorescence intensity and FSC signals in several independent
experiments using different gate settings (see Fig. S3 in the supplemental mate-
rial). Polyphosphate-containing subcommunities were separated by sorting in
order to facilitate their phylogenetic identification. To obtain sufficient DNA for
the generation of 16S rRNA gene clone libraries, at least 106cells were sorted.
(vii) Data evaluation. The acquired data were analyzed as described by
Gu ¨nther et al. (11). The sort gates were placed around particularly abundant or
distinct, rare subcommunities according to similarity in fluorescence and size
properties. The data were determined from two independent experiments ana-
lyzed in two parallels each.
Fluorescence microscopy. To verify reliable staining and purity and to inspect
cell morphologies, the cells were subjected to microscopy and image analysis
(Axioskop [Zeiss] microscope, DXC-9100P camera, and Openlab 3.1.4. [Impro-
vision] software) using light from a 100-W mercury arc lamp. The Zeiss filter set
02 (excitation G 365, BS 395, emission LP 420) was used for examining blue
fluorescence of DAPI-stained cells as well as green fluorescence of TC (0.225
mM)- or DAPI (28 ?M)-stained PAO cells. Red fluorescence from Nile red-
stained cells (PHB detection) was observed using the Zeiss filter set 15 (excita-
tion 546/12, BS 580, emission LP 590). For better visualization of cell proper-
ties, phase-contrast and fluorescence images were merged using the Openlab
software.
TEM. For transmission electron microscopy (TEM), the cells were fixed with
3% glutaraldehyde (Sigma, Germany) in 0.1 M sodium cacodylate buffer (SCB)
(pH 7.2) for 1 h at 20°C. Thereafter the cells were immobilized with 4% agar
(Roth, Germany) in SCB and thoroughly washed four times with SCB. Polyphos-
phate staining was done as described by Jensen (15). Instead of a 20% Pb(NO3)2
solution, a 2% Pb(NO3)2solution was used. Samples were dehydrated in a
graded ethanol series and embedded in epoxy resin (39). Ultrathin sections (80
nm) were transferred to Formvar-coated copper grids and examined without
further staining using an EM 900 transmission electron microscope (Zeiss SMT,
Germany) at an acceleration voltage of 80 kV. Electron micrographs were taken
with a slow-scan camera (Variospeed slow-scan charge-coupled device camera
SM-1k-120; TRS, Germany). The size of the polyphosphate particles was deter-
mined using the iTEM software (Olympus SIS, Germany).
Fluorescence spectra. The fluorescence spectrum of TC hydrochloride (dis-
solved in PBS, 2.25 mM final concentration) was analyzed using the scan mode
of an LS-50B luminescence spectrometer (Perkin-Elmer). The following settings
were used to get information on the fluorescent properties of the compound
when excited at 365 nm, which is the highest excitation wavelength of the
multiline-ultraviolet argon-ion laser of the MoFlo used for single cell analysis:
emission, 300 to 600 nm; number of scans, 2; excitation and emission slit, 10; and
scan speed, 240 nm min?1. The emission spectrum of pure TC without divalent
cations is given in Fig. S2 in the supplemental material. To evaluate the fluores-
cence properties of cells stained with TC, the same instrument settings were
chosen and the front surface accessory option of the Perkin-Elmer device was
used to analyze fluorescence in cell suspensions. Activated sludge cells harvested
from the aerobic cultivation phase and fixed for 1 day were washed twice and
resuspended in PBS to yield an optical density of 0.1. The cells were stained with
0.225 mM TC (final concentration) and analyzed (see Fig. S2 in the supplemental
material). An unstained cell suspension served as a control (see Fig. S2 in the
supplemental material). The peaks at 365 nm are the excitation peaks of the
respective samples.
Pure dissolved TC hydrochloride solution showed a very low fluorescence
maximum at 515 nm, with the respective fluorescence intensity around 0.2.
Unstained cells exhibited a fluorescence peak at between 415 and 420 nm and no
fluorescence above the background level at 515 nm (fluorescence intensity of
about 1.0). TC stained cell suspensions exhibited a small emission shoulder at
420 nm and a large peak with a maximum at 515 nm. The relative fluorescence
intensity of about 10 was 50 times above that of TC solution in PBS and 10 times
higher than the background level of unstained cell suspension.
DNA preparation, 16S rRNA gene cloning, and sequencing. For preparation of
genomic DNA, sorted cells were harvested from the sheath buffer by centrifu-
gation and resuspended in 10 ?l 10 mM Tris-HCl, pH 9.0. Cells were disrupted
by a 10-min microwave treatment at 95°C in a BP-111-RS-IR thermostat micro-
wave device (Microwave Research and Applications Inc.), immediately chilled
on ice for 10 min, and then centrifuged for 10 min at 23,100 ? g and 4°C. From
the supernatant, bacterial 16S rRNA gene fragments were amplified by PCR
using the universal primers 27F and 1492R (22). PCR was performed in a 12.5-?l
reaction mixture containing 6.25 ?l Taq PCR master mix (Qiagen, Germany), 5
pmol of each primer (supplied by Microsynth, Switzerland), and 4 ?l template
DNA with a PTC-200 thermal cycler (MJ Research, USA). For the cycle param-
eters and cloning strategy, see reference 19. Positive clones were screened by
double digestion with the restriction enzymes HaeIII and RsaI (New England
Biolabs, Germany). Partial DNA sequencing of representative clones displaying
different restriction patterns was performed with the BigDye RR Terminator
AmpliTaq FS Kit 1.1 (Applied Biosystems, Germany) and the sequencing prim-
ers 27F and 519R (22). Capillary electrophoresis and data collection were carried
out on an ABI Prism 3100 genetic analyzer (Applied Biosystems). Data were
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Page 4

analyzed with ABI Prism DNA sequencing analysis software, and 16S rRNA
gene sequences were assembled using Sequencher 4.8 (Gene Codes Corp.). The
BLASTN tool (www.ncbi.nlm.nih.gov/BLAST) (3) was used to search for similar
sequences in the GenBank database, and the Seqmatch tool was used to search
for similar sequences compiled by the Ribosomal Database Project (RDP) re-
lease 10.0 beta (http://rdp.cme.msu.edu) (9).
T-RFLP analyses. Bacterial 16S rRNA gene fragments were PCR amplified
with the primers 27F-FAM (labeled at the 5? end with phosphoramidite fluoro-
chrome 5-carboxyfluorescein) and 1492R (22). Labeled oligonucleotides were
purchased from biomers.net (Germany). PCR was performed as described
above. PCR products were purified using the Wizard SV PCR clean-up system
(Promega, Germany) and quantified after agarose gel electrophoresis and
ethidium bromide staining using the GeneTools program (Syngene, United
Kingdom). Purified PCR products were digested with the restriction endonucle-
ase AluI, HaeIII, or Sau3AI (New England Biolabs, Germany). A 10-?l reaction
mixture contained 1 ng DNA (for T-RFLP analyses of single clones) or 20 ng
DNA (for T-RFLP analyses of the entire sample of sorted cells) and 10 units of
restriction enzyme. Samples were incubated at the appropriate temperature for
3 h and then precipitated with sodium acetate (pH 5.5) and ethanol. Dried DNA
samples were resuspended in 20 ?l HiDi formamide containing 1.5% (vol/vol)
GeneScan-500 ROX standard (Applied Biosystems). Samples were denatured at
95°C for 5 min and chilled on ice. The fragments were separated by capillary
electrophoresis on an ABI Prism 3100 genetic analyzer (Applied Biosystems).
The lengths of the fluorescent terminal restriction fragments (T-RF) were de-
termined using the GeneMapper V3.7 software (Applied Biosystems), and their
relative peak areas were determined by dividing the individual T-RF area by the
total area of peaks within the threshold of 35 to 650 bp. Only peaks with relative
fluorescence intensities of at least 20 units were included in the analysis. Theo-
retical T-RF values of the dominant phylotypes represented in the clone library
were calculated using the NEB cutter (http://tools.neb.com/NEBcutter2/index
.php) and confirmed experimentally by T-RFLP analysis using the corresponding
clones as templates. Relative T-RF abundances of representative phylotypes
were determined based on the relative peak areas of the corresponding T-RF.
Nucleotide sequence accession numbers. The 16S rRNA gene sequences de-
termined in this study have been deposited in the GenBank database under
accession numbers EU850352 to EU850394.
RESULTS
Optimization of TC staining of polyphosphate. The TC
staining procedure was optimized with M. phosphovorus NM-1
and a bacterial community derived from activated sludge of the
SBR. Influences of incubation times (0 to 180 min) and TC
concentrations (0.023 to 1.35 mM) on staining results were
tested (see Fig. S4 in the supplemental material). This was
necessary to minimize unspecific staining or dye interactions as
a prerequisite for obtaining quantitative information on indi-
vidual polyphosphate contents. Good results were obtained
after 10 min of exposure to 0.225 mM TC. As the staining was
stable in fixed as well as living samples for 3 h, there was no
sign of efflux pumping of TC.
Polyphosphate staining in pure cultures. Green fluorescent
polyphosphate granules were detected and quantified by flow
cytometry (see below) and fluorescence microscopy (Fig. 1a to
d). Granules were clearly visible in M. phosphovorus, Pseudo-
monas spp., and Paracoccus spp., all harvested after 3 days of
alternating aerobic/anaerobic shifts at the end of an aerobic
phase. The distribution, amount, and size of the polyphosphate
granules varied among the strains as analyzed and observed by
both fluorescence microscopy and TEM (Fig. 1e). Within these
samples the granule diameters varied between 0.25 and 1.33
?m, with average sizes of 0.69 ?m for M. phosphovorus, 0.48
?m for the Pseudomonas sp., and 0.39 ?m for the Paracoccus
sp. (average of 17 granules each). The sizes of the granules in
M. phosphovorus (Fig. 1e) and the Pseudomonas sp. obtained
by fluorescence microscopy were verified by TEM (average of
100 granules each). The granule sizes measured by TEM were
significantly lower. M. phosphovorus showed granule sizes vary-
ing from 0.52 to 0.08 ?m with an average size of 0.26 ?m
(?0.10 ?m), and the Pseudomonas sp. showed granule sizes
varying from 0.17 to 0.04 ?m with an average size of 0.10 ?m
(?0.03 ?m). The differences between results with the two
technologies may partly be an artifact of epifluorescence-based
sizing and the spatial position of the granule within the 80-nm
slices analyzed by TEM.
Samples of M. phosphovorus taken from the aerobic phase
contained 7.02% (?0.17%) green fluorescent cells, whereas
only 1.07% (?0.08%) green fluorescent cells were found in
samples from the anaerobic phase.
Several experiments were conducted to verify reliable, spe-
cific, and quantitative TC staining of cellular polyphosphate.
First,DAPI,whichistraditionallyusedforstainingofpolyphos-
phate granules in bacteria at a concentration range of 18 to 180
?M (see the introduction), was used for comparison. The dye
was applied at a concentration of 28 ?M to stain polyphos-
phate in aerobically grown M. phosphovorus. Flow cytometric
analysis resulted in the detection of 8.34% (?0.32%) and
8.24% (?0.14%) polyphosphate-containing cells after DAPI
and TC staining, respectively. Second, possible unspecific
staining of cellular constituents by TC was tested using Esch-
erichia coli K-12, a bacterium that does not produce polyphos-
phate granules (29). TC (0.225 mM) caused no fluorescence
labeling of E. coli (0.03% ? 0.01%), whereas DAPI (28 ?M)
caused greenish fluorescence of 1.30% (?0.03%) of the
treated cells. Third, Methylobacterium rhodesianum MB126 was
used to test for possible binding of TC to PHB granules. The
presence of PHB was verified using Nile red, which caused
FIG. 1. (a to d) TC-stained cells of M. phosphovorus (a), the
Pseudomonas sp. (b), the Paracoccus sp. (c), and activated sludge (d)
harvested from the aerobic cultivation phase. For improved visualiza-
tion of the polyphosphate granule distribution within the cells, the
fluorescent images were merged with the respective light microscopic
images. Bars, 5 ?m. (e) Cells of M. phosphovorus stained for polypho-
sphates and analyzed by TEM. Bar, 0.2 ?m.
2114 GU ¨NTHER ET AL.APPL. ENVIRON. MICROBIOL.

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bright red fluorescence, whereas TC caused no fluorescence
with PHB (0.03% ? 0.01%). Eventually, the glycogen-accumu-
lating species M. glycogenica, which does not produce polyphos-
phates, was also tested for green unspecific staining, but it
showed no TC fluorescence (0.03% ? 0.01%).
Polyphosphate versus DNA staining in pure cultures. Ana-
lyzing the cytometric proliferation patterns of bacteria gives
information about the growth rate of a species. As we were
interested in the abundance and growth activity of PAOs, we
combined the analysis of polyphosphate contents by TC stain-
ing with the analysis of growth activity based on DNA pattern
analyses using DAPI (1 ?M) by monitoring bacterial chromo-
some numbers. The results are presented in Fig. S5 in the
supplemental material.
To simultaneously obtain information about growth activi-
ties and polyphosphate accumulation of individual cells in a
population, it was important that the DAPI and the TC stains
did not interfere. Indeed, reliable information on DNA pattern
distributions was obtained with cells that had been exposed to
TC prior to the addition of 1 ?M DAPI solution. With this
biphasic staining, it was intended that TC would bind to the
polyphosphate granules before DAPI molecules would bind
there. To detect possible influences of TC on the apparent
distribution of the chromosome numbers, cells were stained
with DAPI alone or in combination with TC. The DAPI fluo-
rescence intensities and distributions of chromosome equiva-
lents (in parentheses) obtained with DAPI only were as fol-
lows: C2n, 12 (50%); C4n, 24 (30%); C8n, 47 (7%); and Cxn, 183
(13%) (Fig. 2a). With DAPI applied in combination with TC,
the results were as follows: C2n, 12 (50%); C4n, 25 (30%); C8n,
47 (7%); and Cxn, 179 (13%). The similarity shows that TC did
not influence quantitative DNA staining with DAPI. The TC-
stained cells (6.1%) were determined after gating and are
shown as white dots in the DAPI-FSC plots (Fig. 2b). The
polyphosphate-containing individuals were distributed over
the entire population, but as a rule, the relative presence of
polyphosphate-accumulating individuals was higher in sub-
populations with more chromosome equivalents. The percent-
age decreased from 28.8% (Cxn) to 4.5% (C8n), 3.3% (C4n),
and 9.3% (C2n). C1ncells were nearly not present within the
sample, which corresponded to a state similar to that shown in
Fig. S5b in the supplemental material, after 6 h cultivation.
When bacterial polyphosphate is stained with high concen-
trations of DAPI (28 ?M), some of the dye also interacts with
the DNA. We therefore checked whether DAPI staining alone
could be used to infer both the DNA and the polyphosphate
contents from the respective blue and yellow-green fluores-
cences. Using the same sample of M. phosphovorus as de-
scribed above (see Fig. S6 in the supplemental material), the
same fraction of polyphosphate-containing cells (6.2%) was
found. Also, the previously observed four subpopulations
based on chromosome equivalents were found and character-
ized by the same mean DAPI fluorescence intensity of each
individual subcommunity as before. However, the apparent
distribution of bacteria between these subcommunities was
shifted toward higher fluorescence intensities as follows: C2n,
13 (5%); C4n, 26 (4%); C8n, 55 (17%); and Cxn, 240 (74%).
These results verified that the linkage of cell proliferation to
polyphosphate synthesis using high DAPI concentrations alone
was not reliable. Therefore, we strongly recommend applica-
tion of the combined labeling approach involving DAPI at low
concentration (1 ?M) and TC (0.225 mM).
PAO dynamics in wastewater communities. The dual-stain-
ing technique was tested by analyzing PAO dynamics in acti-
vated sludge as a proof-of-principle study. The DAPI approach
provides information on subcommunity abundance dynamics
within natural communities, whereas the TC stain highlights
polyphosphate-accumulating community members.
Activated sludge was cultivated in a sequenced batch mode
and subjected to alternating aerobic and anaerobic growth
conditions. Samples were harvested from both cultivation
phases. Biomass was found to be produced at constant rates;
sometimes floc formation was observed after several aerobic/
anaerobic cycles. The MLSS, MLVSS, and ash contents did not
change much during the bioreactor cultivation. We measured
3.0 (? 0.02) g liter?1MLSS, 2.4 (? 0.00) g liter?1MLVSS, and
0.6 (? 0.02) g liter?1ash for the aerobic phase, compared to
2.7 (? 0.58) g liter?1MLSS, 2.2 (? 0.41) g liter?1MLVSS, and
0.5 (? 0.18) g liter?1ash for the anaerobic phase. The nitrate
concentration was close to zero during the anaerobic phases,
indicating nitrate-reducing conditions (see Fig. S7 in the sup-
plemental material). The stationary orthophosphate concen-
tration was set to relatively high values up to nearly 180 mg
liter?1to stimulate the polyphosphate accumulation for con-
venient fluorescence visualization (Fig. 1d). Low phosphate
concentrations as in the OECD medium (e.g., 15 mg liter?1)
led to fewer than 1% PAOs within the community. To visualize
FIG. 2. Pattern of DNA distributions within a DAPI-stained cul-
ture (a) and a DAPI/TC-stained culture (b) of M. phosphovorus.
DAPI-stained cells are shown in black. Prominent subpopulations
characterized by distinct DNA contents are marked with gates (white
ellipses). PAOs within the sample are marked with white dots in the
dot plot and as a white area in the histogram; the percentage of the
PAOs in subpopulations is given on the left side of panel b.
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changed polyphosphate granule accumulation during the an-
aerobic and aerobic phases, cells were harvested 5 min after
entering the anaerobic phase and 105 min after entering the
subsequent aerobic phase (158 mg orthophosphate liter?1in
this sample). They were fixed and stained with DAPI and TC.
As expected, higher quantities of polyphosphate-containing
cells were found during the aerobic cultivation phase (2.9%)
(Fig. 3) than in the anaerobic phase (0.8%).
Within the same sample, up to 19 subcommunities differing
in either polyphosphate or DNA contents as well as cell size
were found. The bars at the bottom of Fig. 3 give their relative
abundances. The numbers and letters in Fig. 3 indicate per-
centages of PAOs within the gates and the denomination of the
subcommunities, respectively. It is remarkable that neither the
number of distinct subcommunities nor the corresponding cell
abundances changed much between the aerobic and anaerobic
phases. However, it was striking that most PAOs were found in
only few of the subcommunities in the sample from the aerobic
phase. These PAOs were characterized by intermediate to high
FSC and intermediate to high DNA contents. We never ob-
served them to be members of the very small and low-DNA-
content subcommunities at the left side of the histogram. The
majority of the PAOs were present in two subcommunities (C
and J), to which they contributed 7.3% and 32%, respectively.
The sizes of the polyphosphate granules varied between 0.39
and 0.41 ?m in diameter (Fig. 1d).
Additionally, the activated sludge cultivation regimen was
modified by altering stationary orthophosphate and carbon
concentrations to prove the effectiveness of the new PAO
detection technique. PAO abundances were followed in de-
pendence on stationary phosphate concentrations, increasing
from 57 mg liter?1(days I/Ia) to 74 mg liter?1(days II/IIa), 97
mg liter?1(days IV/IVa), 151 mg liter?1(days V/Va), and 158
mg liter?1(day III) (Fig. 4) in combination with elevated
carbon contents from 583 mg liter?1(days I/Ia, II/IIa, and III)
to 1,166 mg liter?1(days IV/IVa and V/Va). Samples were
taken in duplicate during the aerobic phases only, on different
days separated by several shifts of the oxygen regimen. As a
result, increased phosphate concentrations clearly led to in-
creased fractions of PAOs within the sludge, from about 0.3%
with 57 mg phosphate liter?1to 4% with 151 mg phosphate
liter?1. Otherwise, the relative abundances of bacterial cells in
the 19 predefined subcommunities remained nearly unchanged
during the cultivation on 583 mg liter?1carbon. Seven major
subcommunities (A, B, C, E, H, L, and N; each above 5%)
comprised nearly 80% of the whole community. When the
carbon concentration was doubled to 1,165 mg liter?1COD, a
different distribution of the community’s individuals devel-
oped. Although the total number of the distinguishable 19
subcommunities remained constant, changed dominance pat-
terns were observed. Now 11 main subcommunities (A, B, C,
D, E, G, H, N, P, S, and T; each above 5%) comprised 90% of
all cells in the 19 subcommunities. This shows the massive
influence of the carbon concentration on the community struc-
FIG. 3. Pattern of DNA distributions (blue, with dominant subcommunities marked with black gates) and PAO distribution (green dots and
numbers within the gates [percent]) of cells harvested from the anaerobic and aerobic cultivation phases and double stained with DAPI and TC.
Subcommunities with a high PAO content are marked with arrows. The amount of cells within each subcommunity is shown in the lower part as
two bars for the respective cultivation phases.
2116 GU ¨NTHER ET AL.APPL. ENVIRON. MICROBIOL.

Page 7

ture. Nevertheless, the general abundances of the key PAO-
containing subcommunities C and J stayed nearly constant
during cultivation on low carbon concentrations, with J slightly
increasing in cell number. A similar stability was observed with
higher carbon concentrations, although the abundance of C
increased from 6 to 9%, whereas cell numbers in J slightly
decreased to 2%. Hence, PAO abundances and dynamics of
key subcommunities were effectively detected, which seems to
depend on stationary orthophosphate concentrations in the
first place, whereas the changing carbon concentrations surely
alter the overall cell abundances within the subcommunities.
Phylogenetic affiliation of PAOs in WWTPs. The reliability
of the dual-labeling technique was also tested by analyzing a
sample harvested from an aeration tank of an industrial
WWTP with an orthophosphate concentration of 15 mg
liter?1. The sample was aerated and kept on ice until flow
cytometric analysis. TC staining revealed that 10.2% of the
cells contained polyphosphate granules. Plotting DNA con-
tents (DAPI) against cells size (FSC) revealed 13 distinct sub-
communities, 8 of which contributed to more than 5% each
and summed up to 85% of all cells in the 13 subcommunities.
The polyphosphate-positive cells were sorted and subjected to
DNA extraction for the phylogenetic identification of PAOs.
A 16S rRNA gene clone library of 78 clones was generated
and screened by amplified rRNA gene restriction analysis.
Based on different restriction patterns, 43 representative
clones were partially sequenced. Phylogenetic affiliations of the
sequences based on the RDP taxonomy and the highest
BLAST hit are given in Table 2. The most frequent phylotype,
comprising 27 clones, was affiliated with the Rhodocyclaceae.
These sequences showed high similarity to clones retrieved
from EBPR sludge, among them those affiliated to the “Can-
didatus Accumulibacter” lineage (13). The second most fre-
quent phylotype, with 13 clones, was assigned to the genus
Pseudomonas. A third group, comprising seven clones, was a
phylotype belonging to the Gammaproteobacteria and related
to glycogen-accumulating organisms retrieved from an EBPR
WWTP (accession no. DQ201885). A phylotype assigned to
the genus Nitrospira was also represented by seven clones. Four
clones were assigned to the genus Tetrasphaera and closely
related to actinobacterial PAOs detected in an EBPR plant
(20). The genus Dechloromonas was also represented by four
FIG. 4. DNA pattern (right panel; labels correspond to those in Fig. 3) and proportions of PAOs (left panel) of an activated sludge community
cultivated in the SBR for 32 days. The cells were harvested from the aerobic cultivation phase and double stained with DAPI and TC. Phosphorous
and carbon concentrations were varied as indicated at the top.
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TABLE 2. Results of sequencing of representative 16S rRNA gene clones
CloneAccession no.Highest BLAST hit (accession no.)/% identity
Taxonomic affiliation
according to RDP
B3 (471)
C10 (475)
C11 (501)
D7 (525)
D10 (483)
E3 (468)
E4 (477)
E6 (458)
E8 (516)
F5 (497)
F11 (485)
H5 (512)
OTUa1
EU850359
EU850366
EU850367
EU850371
EU850372
EU850376
EU850377
EU850379
EU850381
EU850384
EU850389
EU850394
Uncultured betaproteobacterium clone S7 (AF447793)/98
Uncultured betaproteobacterium clone nsc151 (DQ211501)/100
Uncultured bacterium SA34 (AF245349)/99
Uncultured bacterium clone UTFS-002-12-23 (AB166771)/99
Uncultured bacterium clone UTFS-002-12-23 (AB166771)/99
Uncultured bacterium SA34 (AF245349)/99
Uncultured bacterium clone VIR_D5 (EF565151)/99
Uncultured bacterium clone VIR_D5 (EF565151)/99
Uncultured bacterium SA34 (AF245349)/98
Uncultured bacterium clone UTFS-002-12-23 (AB166771)/99
Uncultured bacterium clone VIR_D5 (EF565151)/98
Uncultured bacterium clone D07 (EF589969)/97
27 clones
Rhodocyclaceae
Rhodocyclaceae
Rhodocyclaceae
Rhodocyclaceae
Rhodocyclaceae
Rhodocyclaceae
Rhodocyclaceae
Rhodocyclaceae
Rhodocyclaceae
Rhodocyclaceae
Rhodocyclaceae
Rhodocyclaceae
Rhodocyclaceae
A1 (425)
E5 (443)
E12 (515)
OTU 2
EU850352
EU850378
EU850383
Pseudomonas putida KT2440 (AE015451)/99
Pseudomonas putida isolate 24 (EU438854)/99
Pseudomonas putida BM2 (DQ989291)/99
13 clones
Pseudomonas spp.
Pseudomonas spp.
Pseudomonas spp.
Pseudomonas spp.
B1 (291)
B8 (578)
E2 (463)
OTU 3
EU850358
EU850362
EU850375
Uncultured Nitrospira sp. clone 0B11 (EU499597)/98
Uncultured Nitrospira sp. clone 3 (DQ414437)/99
Uncultured Nitrospira sp. clone 3 (DQ414437)/99
7 clones
Nitrospira spp.
Nitrospira spp.
Nitrospira spp.
Nitrospira spp.
A2 (407) EU850353Uncultured gammaproteobacterium clone GB2917y
(DQ201885)/96
Uncultured gammaproteobacterium clone GB2917y
(DQ201885)/95
Uncultured gammaproteobacterium clone GB2917y
(DQ201885)/94
7 clones
Gammaproteobacteria
A5 (389) EU850355Gammaproteobacteria
A7 (477) EU850357Gammaproteobacteria
OTU 4Gammaproteobacteria
A3 (485)
E9 (451)
F9 (473)
OTU 5
EU850354
EU850382
EU850387
Uncultured eubacterium clone F13.46 (AF495440)/98
Uncultured bacterium clone Ebpr19 (AF255629)/99
Uncultured eubacterium clone F13.46 (AF495440)/98
4 clones
Tetrasphaera spp.
Tetrasphaera spp.
Tetrasphaera spp.
Tetrasphaera spp.
A6 (447)
D11 (504bp)
E7 (535)
OTU 6
EU850356
EU850373
EU850380
Dechloromonas sp. strain A34 (EF632559)/97
Uncultured bacterium clone ORS10C_g11 (EF392932)/99
Dechloromonas sp. strain A34 (EF632559)/97
4 clones
Dechloromonas spp.
Dechloromonas spp.
Dechloromonas spp.
Dechloromonas spp.
B7 (448) EU850361Uncultured actinobacterium clone DOK_NOFERT_clone341
(DQ829293)/96
Uncultured actinobacterium clone DOK_NOFERT_clone341
(DQ829293)/96
2 clones
Actinomycetales
F8 (450)EU850386 Actinomycetales
OTU 7Actinomycetales
E1 (438)
H2 (428)
OTU 8
EU850374
EU850393
Uncultured bacterium clone T015D (AM158382)/91
Uncultured bacterium clone IC-61 (AB255073)/93
2 clones
Candidate division TM7
Candidate division TM7
Candidate division TM7
C12 (530)
G12 (559)
C2 (413)
B10 (532)
D1 (452)
F7 (457)
G10 (449)
B5 (458)
EU850368
EU850392
EU850365
EU850363
EU850369
EU850385
EU850391
EU850360
Uncultured bacterium clone DSSD59 (AY328757)/98
Derxia gummosa (AB089482)/95
Uncultured bacterium clone 197up (AY212650)/99
Ralstonia detusculanense (AF280433)/99
Uncultured bacterium clone TH-141 (AB184983)/98
Uncultured bacterium clone 44 (DQ413103)/99
Uncultured bacterium clone SRRB48 (AB240518)/96
Uncultured Sphingobacteria bacterium clone ADK-SGe02-50
(EF520599)/96
Uncultured bacterium clone HM15 (AM909923)/95
Uncultured bacterium clone LaP15L89 (EF667686)/99
Uncultured bacterium clone mdt16a02 (AY537009)/98
Uncultured bacterium clone 032D06_P_BA_P3 (BX294877)/92
Alphaproteobacteria
Betaproteobacteria
Burkholderiales
Ralstonia spp.
Comamonadaceae
Rhodocyclaceae
Nocardioidaceae
Sphingobacteriales
B11 (522)
F10 (495)
D5 (435)
F12 (574)
EU850364
EU850388
EU850370
EU850390
Acidobacteriaceae
Caldilinea spp.
Bacteria
Bacteria
aOTU, operational taxonomic unit.
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clones. Other phylotypes present in minor proportions were
affiliated with another group of the Actinomycetales (two
clones) and the candidate division TM7 (two clones), and 12
other phylotypes were represented only by single clones (Ta-
ble 2).
T-RFLP analysis revealed that, in contrast to the results
from the clone library, the T-RF assigned to Pseudomonas spp.
was most frequent, comprising more than 50% of the total
peak area (Fig. 5). The Rhodocyclaceae phylotype related to
“Candidatus Accumulibacter” as well as the Tetrasphaera- and
Nitrospira-like phylotypes were also detected in the T-RFLP
profiles but with lower percentages than expected from the
clone library. These results indicate that Pseudomonas spp. are
the predominant polyphosphate-containing PAOs in the
EBPR community analyzed here.
DISCUSSION
Determination and quantification of PAOs in highly diverse
bacterial communities such as activated sludge are of high
interest because of the need for improved phosphorus removal
from domestic wastewater. Scientists and practitioners today
are aware that the community members actually involved in
the biological removal of phosphorus in wastewater are often
unrelated to laboratory strains known to accumulate phos-
phate. Furthermore, many wastewater microorganisms have
the capacity to accumulate the energy-rich compound but do
not do so, for reasons not fully understood. Cultivation-inde-
pendent and quantitative diagnosis of polyphosphate-contain-
ing PAOs is therefore a powerful tool to gain a complete
overview and sound basis for the development of process strat-
egies making the process more reliable.
In this paper the antibiotic TC was proven to show highly
polyphosphate-specific and stable fluorescence. The specificity
for polyphosphate granules was tested with known PAOs such
as M. phosphovorus, the Paracoccus sp., and the Pseudomonas
sp. as well as against a non-phosphate-accumulating strain of
E. coli, the glycogen-producing strain M. glycogenica Lg2T, and
a strongly PHB-accumulating strain of M. rhodesianum. We
compared the new in vivo TC staining method with the tradi-
tional DAPI method and found them to be comparable in
terms of the numbers of labeled cells and the fluorescence
intensities of their phosphate granules. However, the TC stain
proved to be superior to the DAPI stain due to a 15-times-
lower unspecific background labeling. This was regardless of
the fact that the TC fluorescence originates from the strong
chelation of divalent cations, which are present not only in
polyphosphate granules but also in the cell wall, protein com-
plexes, and the cytoplasm (5, 38). Endospores of spore-forming
bacteria such as Bacillus subtilis which do contain calcium ions
also did not show any TC fluorescence above the autofluores-
cence level (not shown). The TC binding to polyphosphate
inside living cells of M. phosphovorus and an activated sludge
community was found to be very stable. Although nearly 40
different efflux systems have been described in various strains
to protect the aminoacyl-tRNA binding sites of bacterial ribo-
somes against the action of TC, we found no sign of TC efflux.
TC molecules strongly chelate with divalent cations. This al-
lows them to enter various bacterial species driven by the cells’
Donnan potential across the outer membrane that acts on the
cations (6). In the periplasm the slightly lipophilic TC mole-
cules are then liberated and diffuse in the cytoplasm.
A very important advantage of TC as opposed to DAPI
staining of polyphosphate was the possibility to combine it with
DNA analysis. Staining of the bacterial DNA gives information
on bacterial growth rates (26, 28) and therefore on activity
states of PAOs when analyzed in combination with TC. DAPI
staining alone of both DNA and polyphosphate granules did
not result in a reliable analysis of the two characteristics. How-
ever, the dual stain with TC and DAPI proved to be a quan-
titative method for PAO detection and DNA content analysis.
We were able to detect polyphosphate-bearing microorgan-
isms even when they contributed less than 1% to a pure culture
or a wastewater community. Since the abundances of PAOs are
of great interest for EBPR processes, we propose the use of
our technique for quick and reliable authentication of PAOs in
WWTPs. Traditional models imply that polyphosphate accu-
mulation occurs during aerobic growth or if the cells are ex-
posed to stress situations such as the decrease of external pH
or carbon substrate concentration (2). However, recent studies
showed that some species still keep polyphosphate granules
even under anaerobic conditions (36). Also, it is now known
that other species which long have served as indicator organ-
isms for polyphosphate accumulation do not reliably accumu-
late polyphosphates under aerobic conditions in the WWTP
process stages (13, 46). For instance, organisms possessing the
ppk gene, which encodes the main enzyme involved in the
synthesis of polyphosphate granules, do not always contain
granules in a WWTP regardless of the oxygen regimen (e.g., E.
coli [30]). Another possible way to approach PAOs in an EBPR
process is the application of oligonucleotide FISH probes spe-
cific for phylogenetic groups that are known as PAOs. How-
ever, these organisms do not always produce polyphosphates
FIG. 5. Relative abundances of T-RF after digestion with the re-
striction endonucleases AluI, HaeIII, and Sau3AI. The T-RF values
were assigned to phylotypes according to the experimentally deter-
mined T-RF values of the respective clones. P, Pseudomonas spp.; R,
Rhodocyclaceae; T, Tetrasphaera spp.; N, Nitrospira spp. With AluI, the
T-RF values of Pseudomonas spp. and Tetrasphaera spp. were identical,
and therefore their relative abundances are summarized.
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and might have such low rRNA contents that they cannot be
detected by conventional FISH techniques. The more sensitive
catalyzed reporter deposition-FISH technique destroys the
structural integrity of the cells, thereby making them unsuit-
able for flow cytometry. Activity-based techniques such as mi-
croautoradiography-FISH have been applied successfully to
detect PAOs (17). However, for routine in situ monitoring of
active PAO populations in industrial WWTPs, our fluores-
cence-based approach using simple and reliable staining tech-
niques and flow cytometry as a quick analytical tool appears
more suitable. Moreover, unlike FISH techniques, it also in-
cludes PAOs of unknown phylogeny.
Within natural communities PAO determination was possi-
ble using the dual-stain technique. Moreover, we found 19
subcommunities according to DNA versus FSC characteristics,
which is an extremely high community resolution when inves-
tigating bacterial communities by flow cytometry. This resolu-
tion may also hint at a highly diverse culture. Additionally, the
DNA/TC patterns revealed information on key PAO subcom-
munity dynamics. It appears justified to conclude that an in-
crease in bacterial cell number within every subcommunity is
caused by cell proliferation and upcoming new cell types (19).
Such an approach based on DNA pattern analysis combined
with the detection of TC-labeled polyphosphates can be re-
garded as highly reliable and quantitative, since there is no risk
of active dye excretion by fixed cells (26, 43).
In fact, the bioreactor experiments combined with the dual-
staining technique revealed high qualitative and quantitative
community stability through aerobic and anaerobic cultivation
modes. This behavior seems to be typical for activated sludge
communities, as previously shown by bulk single-strand con-
formation polymorphism analysis (45). The doubled substrate
concentration had a higher influence and changed the struc-
ture of the community, as was seen by altered abundances in
the 19 subcommunities. It is known that high community di-
versity is a prerequisite for high resilience, as diverse commu-
nities can flexibly react to environmental fluctuations because
of the ecological complementarity of functionally redundant
groups. This has been demonstrated particularly in wastewater
and activated sludge communities (17, 30). Future phyloge-
netic allocation of the upcoming species within the respective
subcommunities can be performed using cell sorting as an
approach which was already reliably applied (19).
Therefore, the application of the dual stain to wastewater
communities enabled differentiation of the entire, complex
community into subcommunities that largely differed in their
growth rates and their polyphosphate contents. The dynamics
of these subcommunities were followed according to cell num-
bers and polyphosphate contents. This was the desired basis for
the separation of the actively polyphosphate-accumulating or-
ganisms from the activated sludge community for phylogenetic
analysis. The results of 16S rRNA gene sequencing verified the
specificity of the staining technique, as the clone library gen-
erated from separated subcommunities displayed a rather low
diversity and was composed basically of phylotypes known as
PAOs, such as the “Candidatus Accumulibacter”-like phylo-
type, members of the genus Pseudomonas, and the Tetraspha-
era-related actinobacteria. Although the clone library probably
does not reflect the complete phylogenetic spectrum present
in the sorted subcommunity, as indicated by several unique
clones, its composition confirms that predominantly PAOs
were captured. Moreover, T-RFLP profiles revealed the
predominance of a single phylotype belonging to the genus
Pseudomonas. Surprisingly, this phylotype was only the second-
most one in the clone library, but T-RFLP profiles recorded
with three different restriction enzymes clearly showed the
predominance of this sequence type. This bias might be ex-
plained by the low coverage of the clone library that led to an
underestimation of the Pseudomonas sequence type. On the
other hand, the relative T-RF peak areas might overestimate
the relative abundances of the corresponding phylotypes, as
sequence types without restriction sites within the 35- to
650-bp threshold are not considered and therefore not in-
cluded in the total peak area. Nevertheless, sequencing and
T-RFLP profiling of 16S rRNA genes verified that PAOs were
specifically detected and captured out of a complex commu-
nity. Thus, the TC dual-staining approach in combination with
high-throughput techniques such as T-RFLP fingerprinting en-
ables the quick and reliable detection and quantification of
distinct PAO groups. In contrast to the FISH technique, it can
also detect PAOs of not-yet-known phylogenetic affiliation.
Our primary goal to quickly detect PAOs and to make them
available for identification as a basis for the improvement of
EBPR processes was therefore achieved.
In summary, we have developed a dual-stain technique to
reliably quantify PAOs and relate them to growth activities
within activated sludge communities in WWTPs. DAPI and TC
are inexpensive dyes that do not change the features of the
cells and can be applied quickly and without complicated sam-
ple handling. The procedure is easy to perform, quick, and
cultivation independent. In the future we aim to combine this
technique with phylogenetic analyses to give an on-line tool for
monitoring EBPR stability and forming a basis for the optimi-
zation of process control.
ACKNOWLEDGMENTS
We thank H. Engewald, C. Su ¨ring, U. Lohse, and S. Jahn for tech-
nical assistance in the laboratory. We thank also W. Geyer for the use
of the fluorescence spectrophotometer and M. Eschenhagen for anal-
ysis of the COD, nitrate, and orthophosphate values of the WWTP
sample.
This work was supported by the German Federal Ministry of Edu-
cation and Research (no. 02WA0700).
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